Visual and Microscopic Evaluation of Streptomyces Developmental Mutants

1. Prepare Instruments, Culture Media, Solutions, and Petri Dishes

1. Autoclave toothpicks in a 50 mL beaker covered with aluminum foil to keep sterile.
2. Sterilize all dry goods that will be used (tips, sticks, glassware, etc.) in an autoclave.
   NOTE: Caution! Follow all safety directions for the autoclave model used. Autoclaves pose a risk for explosions, and burns from steam and contact with hot surfaces, materials, and media. For autoclave safety, view "Proper Use of Autoclaves"³⁴.
3. Prepare the MS agar (Mannitol Soya Flour (SFM))¹⁴ to grow cultures.
   1. Add 5 g of soya flour and 5 g of agar to 1 L flask.
   2. Dissolve 5 g of mannitol in 250 mL of tap water and dispense into the flask containing the soy flour and agar.
   3. Add a foam plug and foil to the opening of the flask and autoclave at 121 °C, 15 psi for at least 30 min.
      CAUTION: This medium easily boils over. Use a flask that can hold at least four times the actual volume of MS agar being autoclaved. A standard pressure cooker can be used in place of an autoclave with similar safety considerations. Alternatively, a microwave oven may be used to sterilize media and materials if access to an autoclave is not possible^35,36. High school teachers may also want to collaborate with local colleges or universities to access autoclaved supplies.
   4. Cool media until comfortable enough to handle. Be careful to not cool it for too long because agar solidifies at temperatures below 50 °C.
   5. Pour MS agar from the flask into sterile Petri dishes until the bottom of each Petri dish is approximately half full (approximately 25 mL of media per Petri dish). Allow to solidify before using agar plates. Store agar plates in a plastic bag at room temperature until needed.
      NOTE: Agar plates may also be stored in a refrigerator, especially if antibiotics are needed in the media.

2. Streak Streptomyces onto Plates for Propagation

1. Streak Streptomyces strains onto MS agar plates to single colonies using a standard method, such as the quadrant streak method as shown in Sanders, 2012³⁷.
   NOTE: An inoculating loop or sterile toothpicks may be used. Optional: Other media are commonly used for Streptomyces species, such as R2YE and minimal media¹⁴.
2. Incubate plates at 30 °C.
3. Restreak plates by picking a single colony every 4–6 days. As colonies age they become prone to random mutation.

3. Create Glycerol Mycelial Stocks for Bacterial Storage

NOTE: Mycelial stocks may be created for all Streptomyces strains, including whi and bld strains, and other developmental mutants that are unable to complete sporulation.

1. Macerate 15–20 colonies in 200–500 µL of 20% glycerol using a sterile wooden dowel applicator.
2. Transfer the slurry of macerated colonies to a 1.5 mL freezer culture tube, using a micropipette or sterile transfer pipette.
   NOTE: The basics of micropipette use are covered elsewhere³⁸.
3. Add enough 20% glycerol to make the final volume 1 mL and gently mix. Store samples at -80 °C.
   NOTE: Caution! Use cryogenic safety gloves and a lab coat to protect skin from contact with extreme cold. Alternatively, a -20 °C freezer may be used with potential reduction in long term storage time. Avoid excessive freezing and thawing to extend viability.

4. Create Glycerol Spore Stocks for Strains Capable of Sporulation

NOTE: Spore stocks are preferable for long-term viability but are only feasible for strains that are capable of completing sporulation.

1. Spread 100 µL of macerated material (from step 3.1) on an agar plate to obtain a confluent lawn of growth³⁷.
   NOTE: Spread plating is shown in Sanders³⁷ (Modification: A turntable is not required.). The plate can be turned with one hand while using the other hand to move the spreader in a gentle back and forth motion across the agar media.
   1. Pipet 100 µL of macerated mycelium to the center of an MS agar plate.
   2. Sterilize a glass or metal spreading tool by partially submerging in 70% ethanol and passing the spreader once slowly through a Bunsen burner flame to speed the evaporation of the ethanol.
      CAUTION: Ethanol is highly flammable. Do not place the flaming spreader into ethanol or allow the flaming drop of ethanol to fall into the ethanol dish. Alternatively, use a disposable sterile cotton swab to spread the bacteria on the plate, which requires no ethanol or flame.
   3. Turn the MS agar plate with one hand, while using a back and forth motion to inoculate the plate with the sterile spreader.
   4. Incubate for 5–7 days at 30 °C until the strain is well-sporulated.
      NOTE: Incubation times will vary depending upon the species and strain.
2. Add 2 mL of sterile saline (0.8% NaCl) to the center of a confluent lawn of Streptomyces cells that has undergone sporulation.
3. Harvest spores by gently rubbing the surface of the mycelium with a sterile inoculating loop (or a cotton bud), slowly moving towards the edge of the lawn.
4. Pipet the saline containing spores into a 15 mL conical tube. Vortex vigorously to break spore chains into individual cells.
5. Centrifuge at room temperature for 5 min at approximately 2,500 x g.
6. Pipet the supernatant and discard. Disperse the spore pellet by vigorous vortex agitation in the small amount of remaining liquid.
7. Resuspend the spores in 1 mL of 20% glycerol by drawing the solution up and down with a pipette.
8. Transfer the glycerol spore stock to a 1.5 mL freezer tube. Store at -80 °C.
   CAUTION: Use cryogenic safety gloves and a lab coat to protect skin from contact with extreme cold. Alternatively, a -20 °C freezer may be used with potential reduction of viability in long term storage time. Avoid excessive freezing and thawing to extend viability.

5. Perform Mutagenesis

1. Perform mutagenesis based on the desired outcomes as referenced in the Introduction and recently reviewed by Baltz, 2016³⁹.
   NOTE: Some phenotyping experiments will not require mutagenesis, such as the intent to identify new species from the environment. Methods of random mutagenesis include the use of a transposon^{8,9,10,11,12,13}, UV radiation, or chemicals¹⁴. Site-directed mutagenesis^{15,16,17,18,19,20} techniques can be used to create specific point mutations, deletions, or other types of mutations.

6. Compare and Record the Visual Appearance

1. Take note of colony morphology for each mutant as compared to the wild-type parent strain after streaking all strains as in step 2. Compare the new isolate to that of a well-studied species, such as S. coelicolor or S. venezuelae, when characterizing new Streptomyces species. Note characteristics (Figure 1) such as basic shape, surface of colony (fuzzy, bald, wrinkled, etc.), opacity, elevation, and pigmentation (distinguish between vegetative mycelium, aerial mycelium, and/or surrounding medium).
2. Label an MS agar plate and streak wild-type and mutant strains in wedge patterns (Figure 1). Be careful that no strain touches another strain to avoid cross contamination. Note the date and time that strains are streaked onto the plate. Incubate the plates at 30 °C.
3. Place grown Petri dishes on colored or white paper to homogenize the background. Write on the paper the strain name, date, incubation temperature, and time from first plate streaking. Take digital picture(s) of the plate with the latter information written on the paper background so that confusion is minimized.
   NOTE: The writing can be cropped from the photograph at a later time if it is to be used for figure preparation.

7. Perform Phase-Contrast Microscopy

1. Sterilize tweezers by washing in 70% ethanol and then passing through a Bunsen burner flame to evaporate the ethanol. Allow to cool.
2. Sterilize coverslips by washing in 70% ethanol and then allowing them to dry in a sterile Petri dish.
3. Prepare a coverslip lift of the bacterial growth to be examined.
   1. Pick up a sterile coverslip with sterile tweezers and place the coverslip on an MS agar plate where bacterial growth is dense. Press on the back of coverslip gently with tweezers to ensure sufficient transfer of bacterial spores and aerial mycelium.
4. Pick up the coverslip from the plate and place it so that the side with the cell material is facing toward the surface of a microscope slide, with a 15 µL drop of 50% glycerol prepared in water for mounting. Reduce air bubbles by placing the coverslip at a 45° angle to the slide and let it fall onto the 50% glycerol.
5. (Optional) Seal with clear nail polish for preservation of the slide.
6. Perform phase-contrast microscopy as described by Frohlich⁴⁰ at various time intervals during the life cycles.
   NOTE: Repeated imaging allows for a more complete analysis and the ability to detect delays in development. Independently repeat the observations to ensure accuracy and reproducibility.
   1. Place the prepared slide on the microscope stage and add a drop of immersion oil to the center of the cover slip. Rotate the 100X phase objective in place and set the condenser turret to the proper "matching" phase setting. Focus the image using only the fine adjustment knob once the objective lens is in contact with the oil.
   2. Examine several fields of view to discern noticeable and consistent differences between mutant strains and the wild type, such as the ability to form spores, spore size and shape, and overall number of spores (see Figure 2).
      NOTE: Alternatives to phase-contrast microscopy include differential interference contrast (DIC) microscopy⁴⁰ and simple staining techniques for use with bright field microscopes⁴¹, such as crystal violet staining.

8. Perform Fluorescence Microscopy

1. Sterilize tweezers by washing in 70% ethanol and then running through a Bunsen burner flame.
2. Pick up a sterile coverslip with sterile tweezers and place on an MS agar plate where bacterial growth is evident. Touch the back of the coverslip gently with the tweezers to ensure sufficient transfer of bacterial spores and aerial filaments.
3. Pick up the coverslip from the plate and place it so that the side with the cell material is facing up, on cardstock for ease in manipulation of the coverslip.
4. Flood the coverslip with ice-cold methanol (~300 µL for a 20 x 20 mm² coverslip) and let it completely air dry.
   CAUTION: Methanol is a mutagen. Avoid direct contact with skin.
5. Wash the coverslip three times in Phosphate Buffered Saline (PBS)⁴² by gently dispensing and removing the solution.
6. Add 15 µL of 100 μg/ml Propidium Iodide and/or 10 μg/mL Wheat-Germ-Agglutinin conjugated to fluorescein isothiocyanate (WGA-FITC) made in 50% glycerol to a labeled microscope slide. 
   NOTE: Propidium iodide stains DNA red to detect chromosomal DNA location while WGA-FITC stains the cell wall green.
7. Invert coverslip face down onto the drop of fluorescent stain on the glass slide. (Optionally) seal with clear nail polish for preservation of slide.  As the dyes are light-sensitive, incubate for 15 minutes in a darkened or dimly lit room.
8. Observe the slides under a 100X objective lens using a phase-contrast or DIC microscope equipped with epifluorescence excitation cubes for propidium iodide ('Texas Red' Filter Set or 535/617 nm excitation/emission maxima) and FITC (FITC filter set or 490/525 nm excitation/emission maxima).
9. Place the slide on the stage and focus using the coarse and fine adjustment knobs.
10. Observe the fluorescence as shown elsewhere⁴³.
11. Use a free image analysis software package that can help in the identification of spores with lower fluorescence intensity⁴⁴.